Oleosome interfacial engineering to enhance their functionality in foods

This study aimed to increase the physical stability of native sunflower oleosomes to expand their range of applications in food. The first objective was to increase the stability and functionality of oleosomes to lower pH since most food products require a pH of 5.5 or lower for microbial stability. Native sunflower oleosomes had a pI of 6.2. One particularly effective strategy for long-term stabilization, both physical and microbial, was the addition of 40% (w/w) glycerol to the oleosomes plus homogenization, which decreased the pI to 5.3 as well as decreasing oleosome size, narrowing the size distribution and increasing colloidal stability. Interfacial engineering of oleosomes by coating them with lecithin and the polysaccharides xanthan and gellan, effectively increased stability, and lowered their pI to 3.0 for lecithin and lower than 3.0 for xanthan. Coating oleosomes also caused a greater absolute value of the ζ-potential; for example, this amount was shifted to −20 mV at pH 4.0 for xanthan and to −28 mV at pH 4.0 for lecithin, which provides electrostatic stabilization. Polysaccharides also provide steric stabilization, which is superior. A significant increase in the diameter of coated oleosomes was observed with lecithin, xanthan and gellan. The oleosome sample with 40% glycerol showed high storage stability at 4 °C (over three months). The addition of glycerol also decreased the water activity of the oleosome suspension to 0.85, which could prevent microbial growth.


Introduction
In eukaryotes, oleosomes are naturally emulsified oil droplets containing triacylglycerols (TAGs) in the core (94-98% w/w), covered and stabilized by a unique protein/phospholipid membrane layer. This unique structure makes them form a natural oil-in-water suspension which serves as energy stores for the germination and growth of seedlings (Guzha et al., 2022 andHuang, 1992). Isolated oleosomes from different sources have a spherical shape and possess diameters ranging from 0.2 to 2.5 μm controlled by the relative TAGs to the oleosin (structural proteins found on oleosome surface) ratio (Nikiforidis, 2019). However, previous studies showed oleosome extraction method greatly affects their particle size, ranging from nanoscale to microscale (Nikiforidis et al., 2013). The oleosome membrane that consists of mainly phosphatidylcholine (PC) and phosphatidylserine (PS) monolayer (about 2% w/w of oleosome) that are negatively charged interact through electrostatic attractive forces with the basic amino acid residues of surface proteins (oleosins, caleosins, and steroleosins), whose content is about 0.6-3.0% w/w (Tzen et al., 1993;Shimada and Hara-Nishimura, 2010;Deleu et al., 2010). Among surface proteins, oleosins are the main proteins which are small alkaline proteins with a molecular mass ranging from 15 to 26 kDa (Tzen and Huang, 1992). Amino acid sequence analysis of oleosin has shown the existence of three structural domains, including an amphipathic N-terminal domain, a central hydrophobic hairpin structure domain that is pinned in the TAG core, and an amphipathic α-helical domain near the C-terminus. These secondary structures cause the protein to reside stably on the phospholipid monolayer on the surface of the oleosomes (Jolivet et al., 2004). While caleosins have a shorter hydrophobic domain similar to oleosins (Frandsen et al., 2001), it contains an N-terminal region that is hydrophilic and contains a single Ca 2+ binding site exposed to the aqueous phase (Frandsen et al., 1996). Subsequently, the aqueous phase's polarity significantly influences the secondary structure of caleosins, and their properties might change in different environments (Purkrtova et al., 2007). Steroleosins are relatively bigger proteins (above 35 kDa) with a hydrophobic anchoring segment at the same size as caleosins and a hydrophilic domain in contact with an aqueous phase (Lin et al., 2002).
Oleosomes as natural emulsion potentially have many applications in foods such as sauces, salad dressing, coffee whitener, mayonnaise, milk cream alternatives, yogurt, and ice cream, and can be incorporated into plant proteins and polysaccharides for making meat alternatives, plant-based cheese, and pastry (Nikiforidis et al., 2014). Aqueous extraction of crushed hydrated oilseeds yields both oleosomes and water-soluble proteins in water, which is usually followed by filtration and centrifugation to cocentrate oleosomes. The oleosome extraction parameters, including the degree of grinding, oilseed/water ratio, extraction time, and temperature, significantly affect the extraction yield (Rhee et al., 1972). After centrifugation, up to 75%-90% (w/w) aqueous suspension of oleosome (cream layer) can be separated (Nikiforidis and Kiosseoglou, 2010;Nikiforidis et al., 2014).
The low chemical and microbial stability of oleosomes is the main barrier to their use in food products (Chang et al., 2013). The chemical stability of the oleosome suspension and its flocculation is mainly dependent on electrostatic attraction and repulsion interactions between oleosome-associated surface proteins (Iwanaga et al., 2008). The charge properties of the oleosomes' surface can be a major factor contributing to the stability of the oleosomes, and changing pH can have a large effect on their stability. Previous studies showed that the isoelectric point (pI) of oleosomes ranges from 5.7 to 6.6, with generally a negative surface charge at neutral pH (Maurer et al., 2013;Garcia et al., 2021). If the pH of the oleosome suspension drops below their pI, they start aggregating because of the relatively weak electrostatic repulsion forces operating between the droplets close to the pI (Sukhotu et al., 2014). This phenomenon severely limits their potential applications in foods.
In this study, we attempted to deposit natural surface-active components such as lecithin and polysaccharides on the oleosome surface to create a multilamellar structure surrounding the oil droplet and provide steric and electrostatic barriers against flocculation and enhance mechanical strength. Multi-lamellar "structured" oleosomes, or coated oil bodies, will have enhanced stability and improved mechanical properties. Moreover, the addition of glycerol to decrease the oleosome suspension's water activity and increase their storage stability will be discussed. The new structured oleosomes could potentially be added to spreads, cream, and processed cheese at acidic pHs, as well as better withstand processing operations such as pasteurization and homogenization.

Coating oleosomes with lecithin, polysaccharides and glycerol
To understand the effects of adding glycerol, lecithin, and polysaccharides on increasing the physical stability of oleosomes, a mixture of 40% (w/w) glycerol, 40% (w/w) oleosomes, and 20% (w/w) buffer (25 mM NaHCO 3 ) was prepared and homogenized at 15,000 RPM (IKA magic LAB, Wilmington, NC, USA) for 1 min. In comparison, coated oleosomes with lecithin were prepared by adding 0.05%, 0.1%, 2%, 3%, and 10% (w/w) lecithin and mixed with oleosomes using a magnetic stirrer for an hour at room temperature, followed by homogenization at 15,000 RPM for 1 min. To study the effects of coating oleosomes with hydrocolloid gums on increasing their colloidal stability, we first tested the solubility of 0.1% (w/w) gums (carrageenan, pectin, fenugreek, locust bean, guar, gellan, and xanthan) in water. Pectin, carrageenan, and xanthan showed high solubility in water at room temperature, while the solubility of Fenugreek, guar, and gellan gums in water was quite low. Heating above room temperature and mixing were necessary to solubilize fenugreek, guar, and gellan gums in water. Gellan was solubilized in water only after 30 min of heating at 90 • C.
In the second step, after completely dissolving gums in water, the pH of 0.1% (w/w) gum solutions in water was decreased to 4.0 using 0.1M HCl. Then oleosomes were added to each gum solution and mixed for 30 min to obtain final concentrations of 10% (w/w) oleosomes in the gum dispersions, followed by homogenization at 15,000 RPM for 1 min.

Particle size analysis
The particle size distribution of the oleosomes was evaluated using a static multi-angle light scattering (Mastersizer 2000; Malvern Instruments Co., Ltd., Worcestershire, UK). The oleosome samples were diluted with sodium carbonate buffer solution (25 mM NaHCO 3 ) in different concentrations to a pH of 8.3. The relative refractive index of oleosome suspensions and the continuous water phase was 1.474 and 1.330, respectively. The results were reported as the volume surface mean diameter (D[3,2]) and volume-weighted mean diameter (D[4,3]).

ζ-potential analysis
The ζ-potentials of the oleosomes were determined using a ζ-potential analyzer (Malvern Instruments, Malvern, UK). The oleosome suspension (70% w/w) was diluted in water to the concentration of 0.001% (w/v). The pH of the diluted suspension was set in the range of 3.0-9.0 using hydrochloride acid (0.1M) or sodium hydroxide (0.1 M). Then after, the ζ-potential of samples was determined at 25 • C.

Melting point analysis
A differential scanning calorimetry (DSC) model TA Q2000 (TA Instruments, Mississauga, ON, Canada) was used to determine the melting points of oleosome suspension (70% w/w). Nitrogen was used to purge the system at 18 mL/min flow rate. This study determined the melting point of 70% (w/w) oleosome by heating samples from 20 • C to 80 • C at the heating rate of 5 • C min − 1 .

Static surface tension measurement
Static surface tension measurements were conducted at the air-water interface using a Sigma force tensiometer (Biolin Scientific, Linthicum Heights, MD, United States) and a DuNuoy ring. Measurements were conducted immediately after transferring samples. The surface tension provided was estimated using the equation of γ=(F/4πRf). Where F is the maximum force measured when pulling the Du Nuoy ring out of the water phase and overcome surface tension, R is the average radius of the Du Nuoy ring used, and f is the Huh and Mason correction factor calculated as f = R/r (R = radius of the Du Nuoy ring and r = radius of the wire).

Compression isotherms
Compression isotherms of samples were measured at the air-water interface using a Kibron Microtrough G1 Langmuir-Blodgett trough (Kibron, Sweden), controlled using KBN LayerXPro software (Kibron, Sweden). After transferring samples in the trough, interfacial films were immediately compressed from 16,500 mm 2 to 1650 mm 2 , using mobile barriers moving at a speed of 30 mm/min. The pressure was monitored using a Wilhelmy plate during each compression.

Confocal laser scanning microscopy (CLSM)
The CLSM analysis of oleosomes and their potential aggregation was studied visually using an inverted confocal laser scanning microscope Leica DM IRBE (Leica Microsystems, Heidelberg GmbH, Germany) and a Leica TCS SP2 (Leica Microsystems, Heidelberg GmbH, Germany). After dilution in sodium carbonate buffer solution (5% w/v), the oleosome samples were dispersed on a microscope slide (5 μL). Then the prepared oleosome suspensions were stained (10% v/v) with different dyes (Nile Blue (0.01% w/v, and Rhodamine B (0.05% w/v). After covering the microscope slides with a coverslip, it was fixed using nail polish on the corners of the coverslip. Microscope slides were stored overnight to penetrate the dye in the oleosome structure better. Ar/Kr and He/Ne lasers were operated at excitation wavelengths of Nile Blue, and Rhodamine B at 488 nm, and 510 nm, respectively. The confocal images were recorded at a magnification of 20X and 63X.

Oleosome polysaccharide complex suspension stability test
In this analysis, suspension of oleosome in aqueous solutions containing 0.1% (w/w) of gellan, or 0.1% (w/w) of xanthan that already set their pH at 4.0 using HCL (0.1M) was prepared (final concentration of 10% w/w). After adding oleosomes in the gum solutions, they were mixed for 30 min and the oleosome-polysaccharide mixtures were homogenized by passing through a rotostator homogenizer at 15000 RPM for 1 min. To compare their stability (phase separation as a function of time), the homogenized oleosome-polysaccharide mixtures were stored at 30 • C for 24 h.

Light microscopy
The morphology of oleosomes was studied using an OMAX light microscope (China). The diluted oleosome suspension (10% w/w) was pipetted on a microscope glass slide and coved with a glass cover. The oil bodies were observed with 20X and 40X optics.

Rheological analysis
The rheological properties of the oleosome suspensions were analyzed using both dynamic and static rheological properties. For this purpose, an Anton Paar MCR 302 rheometer (Saint-Laurent, Quebec, Canada) with a cuvette cell geometry (concentric cylinder) was used to test the following parameters.

Amplitude sweep test
In this analysis, the shear strain (oscillating) was a logarithmic profile ramp with initial at 60% and final at 200%. For angular frequency, the constant profile set at 1 S − 1 . The analysis temperature was set at 20 • C.

Frequency sweep test
For the dynamic frequency sweep measurement, the shear strain (oscillating) was constant at 100% value and angular frequency profile was ramp logarithmic, initial at 0.1 S -1 and finalized at 200 s − 1 . The analysis temperature was set at 20 • C.

Flow curve analysis
In this analysis, the shear rate was set on a ramp linear profile, starting at 10 S − 1 to 100 S − 1 . The analysis temperature was set at 20 • C.

The effect of salt (NaCl) on oleosome surface charge
In this experiment, first different concentrations (0 M, 0.01 M, 0.02 M, 0.05 M, and 0.1 M) of sodium chloride in de-ionized (DI) water were prepared. Then a 10% oleosome in DI water was mixed for 10 min and homogenized using a rotostator at 15,000 RPM for 1 min. The final pH of the suspension was 8.14. In the next step, 0.25 ml of 10% oleosome suspension was added to each saline solution and mixed for 1 min. The final pH of saline solutions was 6.2. In the final step, the ζ-potentials of the oleosomes in saline solutions were determined using a ζ-potential analyzer.

Water activity analysis
The water activity analysis was performed using an Aqua Lab water activity analyzer model 4 TEV (Decagon Pullman, WA, USA) with the temperature control. The analysis temperature was set at 21 • C. To calibrate the water activity analyzer, two standard solutions were used including a 6 M sodium chloride in water (aw = 0.76), and a 13.41 M lithium chloride in water (aw = 0.25).

Statistical analysis
All experiments were carried out in triplicate, and the results are presented as average ± standard deviation. Statistical analysis was carried out using GraphPad Prism software version 5.0 (La Jolla, CA, USA). All analyses were run in duplicates and results were stated as mean values ± standard deviations. Data were evaluated using one-way ANOVA and probability of p<0.05 was considered as significant.

Evaluation of oleosome suspension
In order to determine the initial quality characteristics of our sunflower olesome suspensions (70% w/w oil), the pH, particle size distribution, ζ-potential, and microscrostructure were evaluated. The results showed the pH of the commercial olesome samples was 7.79 ± 0.05, and the oleosomes were homogenously dispersed in water and no aggregation was observed. The average diameter of the oleosomes was d4,3 (volume-mean diameter): 5.12 ± 0.44 μm and d3,2 (Sauter-average diameter): 3.83 ± 0.62 μm (Fig. 1). The ζ-potential of oleosomes at pH 7.8 was − 42.4 ± 2.1 mV. The morphology of oleosomes is shown in Fig. 2.
To better understand the influence of pH and ionic strength on the colloidal stability of sunflower oleosomes in water, the particle size distribution and ζ-potential were measured in different pH and ionic strengths (Fig. 3.).
ζ-potential analysis was carried out to determine the surface charge of the oleosomes at different pH value in the range [4-10]. Surface charge is a critical factor affecting the colloidal stability of oleosome suspensions. The ζ-potential analysis showed that the pH stability of oleosomes was very poor, with an isoelectric point of ~6.2. Fig. 3 (left) shows the change in ζ-potential as a function of pH for sunflower oleosome suspension (10% w/w) in water. The colloidal stability of oleosomes and the rate of flocculation are mainly dependent on the electrostatic attraction and repulsion interactions between oleosomeassociated proteins (Iwanaga et al., 2008). The ζ-potential of oleosomes ranged from +40 mV at pH 4.0 to − 42.0 mV at pH 10.0, with an isoelectric point of approximately 6.2 (Fig. 3, left). So, at lower pH, protonation of the oleosome-associated proteins results in an increase in positive charge, while at high pH, deprotonation takes place, resulting in negatively charged oleosomes. This suggests that oleosomes will destabilize at a pH very close to neutrality and definitely at acidic pHs. Fig. 4 shows the flocculation of a 10% (w/w) oleosome suspension in water at pH 6.5.
The change in the ζ-potential of oleosomes at different ionic strengths is shown in Fig. 3, right. An increase in NaCl concentration resulted in a significant decrease in the ζ-potential of the oleosomes. After the addition of NaCl, the magnitude of the ζ-potential was increased from − 30 mV to 0 mV at 100 mM NaCl concentration. In general, increasing the ionic strength of the solution causes a reduction of the charge density of colloidal suspensions due to electrostatic screening effects (McClements, 2016). The negative charge of oleosomes is mainly attributed to surface proteins (mainly oleosins) and phospholipids interface, and small changes in the ionic strength of the solution can cause a decrease in the negative charge density of the oleosome interface. Similar results have been reported for the other types of oleosomes, including maize, soybean, and safflower, and a significant reduction in ζ-potential was observed with the addition of salt (Sukhotu et al., 2014;Iwanaga et al., 2007). The aggregation of oleosomes and coalescence that follows at low pH and high salinity will severely limit the application of oleosomes as food ingredients in many food products (Nikiforidis et al., 2014).

Effect of adding glycerol on oleosome stability
Changes in the particle size distribution of the oleosome mixture in glycerol were monitored during the three-month storage of the mixture at 4 • C. The results showed during the three-month storage of sunflower oleosomes in glycerol, no drastic change in the average particle size of oleosomes (Fig. 5). Homogenization of oleosomes with glycerol caused a significant reduction in the average diameter of oleosomes from 5.12 ± 0.44 μm to 2.63 ± 0.16 μm.
Addition of glycerol to oleosome suspensions (Fig. 7) on changes of ζ-potential at different pH (3-9) showed the pI of suspension was decreased from 6.3 for sunflower oleosomes to 5.3 in the mixture of oleosomes and glycerol (Fig. 6). This demonstrates that glycerol addition and homogenization decreases both pI and size of oleosomes that generally leads to increase of colloidal stability.
The addition of glycerol to oleosome suspension also led to a decrease in the water activity of glycerol/oleosome mixture from 0.993 ± 0.001 to 0.856 ± 0.002, which potentially can limit microbial growth. Glycerol has been extensively used to control water activity in food and pharma (Marcolli and Peter, 2005).

Coating oleosomes with phospholipids
The main purpose of adding phospholipids to the oleosome  suspension was to improve the mechanical and chemical stability of the oleosomes. The effect of the addition of 0.2% (w/w) lecithin on oleosome particle size is shown in Table 1. Statistical analysis (unpaired t-test with Welch's correction) of the meandifferences in particle size distribution showed a significant increase in the diameter of coated oleosomes for 10% oleosomes with 0.2% lecithin. For example, based on increases in the D [4,3] value in Table 1, we estimate a 447 nm deposition on the oleosome surface. We have thus effectively created a "coated" oleosome with properties that significantly differ from the original one. We proceeded then to generate binding curves (Fig. 8). Based on the obtained results, it is clear that the addition of lecithin to oleosomes lead to an increase in oleosome size.
The deposition of lecithin on the oleosome surface caused a dramatic shift in the pI to lower pH values. Fig. 9 shows the ζ-potential vs. pH profile for coated oleosomes with 1% lecithin. This suggests that coated, or structured, oleosomes will withstand much better acidic conditions. Therefore, their colloidal stability is enhanced, and potentially their usability in food products can be improved.
Since rhodamine B forms adducts with phospholipids and does not require a heat treatment, it was used to stain lecithin-coated oleosomes (Castro et al., 2012). The light micrograph image showed a more prominent surface staining of the lecithin-coated oleosomes (dark-red circles), corroborating the finding that oleosomes are coated with the sunflower lecithin (Fig. 10).
The compare the physical stability between coated oleosomes with lecithin and oleosomes without lecithin at low pH and high shear, a 50% oleosome sample was mixed with 1% (w/w) sunflower lecithin at pH 5.0 and homogenized at 15,000 RMP for 1 min using a rotostator. The samples were then stored at 4 • C for five days. A greater physical stability of lecithin-coated oleosomes relatvive to uncoated homogenized oleosomes was observed (Supplement Fig. 1).
Since the oleosome surface is permeable to hydrophobic molecules, this ability can be exploited to stain the oleosomes with hydrophobic dyes like Nile Blue and visualize them under confocal light (Karefyllakis et al., 2019). Confocal Laser Scanning Microscope (CLSM) analysis of uncoated oleasomes and coated oleosomes with lecithin stained red by Nile Blue are shown in Fig. 12. In these selected images, the multilayer coating of lecithin on the oleosome surface can be observed (Fig. 11b).
In the next part of this study, the rheological characteristics of coated oleosome with lecithin were compared to uncoated oleosomes. For this Fig. 5. Change in the particle size distribution of 40% (w/w) sunflower oleosome, and 40% (w/w) glycerol mixture in 20% (w/w) sodium bicarbonate buffer (20% w/w) during storage at 4 • C. Fig. 6. Changes in ζ-potential as a function of pH for a (left) natural oleosomes in sodium bicarbonate buffer and (right) mixture of 40% (w/w) oleosome and 40% (w/w) glycerol in 20% (w/w) sodium bicarbonate buffer.  purpose, a traditional flow curve (viscosity vs. shear rate) of samples showed an unstructured liquid with a classic Newtonian behavior, and a constant viscosity as a function of shear rate (Fig. 12). Upon addition of 1% lecithin to 50% oleosome, the viscosity of the oleosomes increased form 6.9 mPa s to 9.4 mPa s (@ 40s − 1 ). This suggests a larger particle size or some oleosome-oleosome interactions. Interestingly, the viscosity of the liposomes was only 2.15 mPa s. Thus, the observed effect is clearly due to the coating of the oleosomes. The average viscosity of samples at different shear rates is shown in Table 2.
The results of dynamic oscillatory rheological measurements, namely as an amplitude sweep and a frequency sweep, are shown in Figs. 13 and 14, respectively. The amplitude sweeps (Fig. 14) clearly show the fact that the material has very weak structure, and the loss and storage moduli are very small. Moreover, the G", or loss modulus, is greater than the storage modulus (G'), clearly suggesting coated and uncoated oleasomes are, in fact, liquids with very little to no elastic character.
In Fig. 14, frequency sweeps of samples show a strong frequency dependence for both moduli in all samples, and the loss modulus was always greater than the storage modulus. This is characteristic of fluids with no particular structure. The power-law exponent for all samples in the frequency sweep was ~1.25.

Coating of oleosomes with polysaccharides
Previous studies showed attractive electrostatic interactions formed by protein-hydrocolloids complex interactions could potentially be used to improve the physical stability of oil-in-water emulsions (Zambrano and Vilgis, 2023;Moreau et al., 2003;Guzey and McClements, 2007;Mert and Vilgis, 2021).
Addition of oleosomes to the pH 4.0 gum dispersions increased the pH to different extents depending on the gum. (Table 3). The final pH for 10% (w/w) sunflower oleosome suspension in 1% (w/w) pectin was lower than the pI of sunflower oleosomes (6.2), while for fenugreek, gellan, and xanthan, the final pH was close to the pI of the oleosomes. A higher final pH (7.2) was obtained for sunflower oleosome suspensions in 0.1% carrageenan, locust bean, and guar gums. In the final step, the physical stability of 10% oleosomes in 0.1% gum complex suspensions was compared after storing samples at 30 • C for 24 h.
The result of physical stability is shown in Fig. 15. Colloidal instability was observed for carrageenan, pectin, fenugreek, locust bean, and guar systems, while no phase separation was observed for 10%, sunflower oleosomes coated with 0.1% xanthan or 0.1% gellan hydrocolloids.
Xanthan and gellan are anionic polysaccharides with good temperature, pH, and ionic strength stability. Based on our colloidal stability test (Fig. 15), we conclude that xanthan and gellan as anionic polysaccharides can be adsorbed onto the surfaces of oleosomes at pH values   where the oleosomes have a net positive charge on their surface (Gu et al., 2005;Mert and Vilgis, 2021). The addition of 0.1% (w/w) xanthan and gellan gums to 10% oleosomes caused a change in the surface charge of oleosomes. It is generally accepted that the greater the absolute value of the ζ-potential, the stronger the electrostatic repulsive force and the more stable the emulsion. At their pI, oleosomes have poor stability due to aggregation. So, the stability of oleosomes towards aggregation can be improved by coating them with polysaccharides (xanthan and gellan) that form thick charged interfacial layers that increase both electrostatic and steric repulsion between oleosomes. After adding oleosomes to the xanthan and gellan solutions, the pH of the suspension was increased close to the pI of oleosome surface proteins. This could be the reason for the high affinity of xanthan and gellan for the surface proteins on the oleosomes compared to lecithin. In this study, an oleosome complex suspension containing 0.1% xanthan or 0.1% gellan showed a large negative surface charge in the pH range studied, from − 60mV at a pH of 8.0 to − 13mV at a pH of 3.8 (Fig. 16).
The rheological properties of 10% oleosomes compared to coated oleosomes with 0.1% xanthan were also studied to better understand the effect of adding 0.1% xanthan on viscosity and dynamic shear rheology of the suspensions. Results are shown in Fig. 17. Coated 10% oleosomes with 0.1% xanthan complex suspensions were pseudoplastic (Fig. 17a), and addition of xanthan gum caused an increase in the viscosity of the suspension, which can also affect the stability against separation. The G ′ and G" of 10% oleosome/0.1% xanthan complex suspension were very small, and the amplitude sweep was unremarkable, with the expected G">G' observed (Fig. 17b). The zero-shear viscosity for this sample was 135 ± 1.79 mPa s from a power-law fit to the data. While the viscosity of oleosomes at the shear rate of 10 S − 1 was 7.74 mPa s, after coating oleosomes with xanthan the viscosity at the same shear rate was increased to 13.16 mPa s.

Static surface tension and compression isotherms of oleosomes
To evaluate the effects of coating 0.12% oleosomes with lecithin and xanthan, oleosomes were mixed with 0.0024% (w/w) lecithin or 0.00024% (w/w) xanthan on static surface tension and compressional behavior, three samples were prepared and were analyzed either immediately or after one day of aging (the pH of the suspension was adjusted to pH = 7.2, using 0.1 M NaOH solution).
The results of these analyses showed that oleosomes were the most interfacially active among fresh samples and formed the films softest upon compression. (Table 4). The compression isotherm results are shown in Fig. 18 Addition of either lecithin or xanthan caused a higher compressional modulus than uncoated oleosomes, suggesting a higher mechanical strength of the oleosomes upon coating with either lecithin  or xanthan. Both lecithin and xanthan coating increased the surface tension of the oleosome suspension, indicative of changes in the nature of the oleosome surface and indirectly the surface tension of the suspension. It would seem the surface of the oleosomes had become more hydrophilic upon being coated with xanthan and lecithin.

Conclusion
In this study, the interface of oleosomes was engineered to increase their physical stability. Deoiled lecithin and polysaccharides such as xanthan and gellan, as well as plain glycerol addition were studied. Results showed that the coating of oleosomes with lecithin and xanthan plus homogenization effectively decreased particle size, increased thermal stability, and lowered their pI. Moreover, a mixture of 40% glycerol+40% oleosome+20% buffer showed a high storage stability at 4 • C, over 3 months. The addition of glycerol also decreased the water activity of the oleosome suspension to 0.85, which potentially could prevent microbial growth.  Frequency sweeps (rad/s) at a shear strain of 100% for (a) 1% lecithin liposomes, (b) 50% uncoated oleosomes, and (c) 50% oleosomes coated with 1% lecithin.

Table 3
Change in pH of 0.1% aqueous solutions of gums before setting to pH 4.0 and after adding oleosomes.

Declaration of competing interest
The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: Alejandro Marangoni reports financial support was partially provided by the Natural Sciences and Engineering Research Council of Canada and partially by Botaneco, Inc. Hargreaves, Mata and Guldiken are    employees of Botaneco, Inc., a company that produces oleosomes for commercial purposes.

Data availability
Data will be made available on request.